Biotin- and Glycoprotein-Coated Microspheres: Potential Surrogates

Article pubs.acs.org/est

Biotin- and Glycoprotein-Coated Microspheres: Potential Surrogates for Studying Filtration of Cryptosporidium parvum in Porous Media Liping Pang,*,† Urszula Nowostawska,‡ Louise Weaver,† Gabrielle Hoffman,† Anjuman Karmacharya,† Alexandra Skinner,† and Naveena Karki† †

Institute of Environmental Science and Research Ltd., PO Box 29181, Christchurch, New Zealand Chemistry Department, University of Otago, PO Box 56, Dunedin, New Zealand

‡

ABSTRACT: Cryptosporidium parvum is a waterborne pathogen, yet no suitable surrogate has been established for quantifying its filtration removal in porous media. Carboxyl polystyrene microspheres with size, density, and shape similar to C. parvum were coated with biotin (free and containing amine, NH2) and glycoprotein. These biomolecules have isoelectric points similar to C. parvum (pH ≈ 2), and glycoprotein is a major type of surface protein that oocysts possess. Zeta potential (ζ) and filtration removal of particles in sand of two different grain sizes were examined. Compared to unmodified microspheres, modified microspheres achieved a superior match to the oocysts in ζ, concentration, mass recovery, and collision coefficient. They showed the same log reduction in concentration as oocysts, whereas results from unmodified microspheres deviated by 1 order of magnitude. Of the three types of modified microspheres, glycoprotein-coated microspheres best resembled oocyst concentration, despite having ζ similar to NH2-biotin-coated microspheres, suggesting that surface protein also played an important role in particle attachment on solid surfaces. With further validation in environmental conditions, the surrogates developed here could be a cost-effective new tool for assessing oocyst filtration in porous media, for example, to evaluate the performance of sand filters in water and wastewater treatment, water recycling through riverbank filtration, and aquifer recharge.

1. INTRODUCTION Cryptosporidium parvum (C. parvum) is a waterborne protozoan that causes gastrointestinal illness.1 The oocysts of C. parvum are shed in the feces of infected humans and animals, particularly young animals. The infectivity of C. parvum oocysts is relatively high and ingestion of fewer than 10 oocysts can lead to infection.2 C. parvum oocysts are commonly found in surface waters and have been detected in some drinking water supplies.2,3 Because of their thick cell wall, C. parvum oocysts are extremely resistant to disinfection.2 Inadequate removal of C. parvum in water treatment plants and groundwater has resulted in many cryptosporidiosis outbreaks.4,5 However, C. parvum is not routinely monitored during water and wastewater treatment because enumeration of oocysts is very expensive and labor intensive. No quantitative surrogates for oocyst removal during water treatment have been identified, and traditionally, filtration removal of oocysts in water treatment has been assessed by turbidity.6 Because of its resistance to standard disinfection, C. parvum removal in water and wastewater relies largely on filtration through porous media. Consuderable effort has been made to understand the process of oocyst removal by filtration as reflected in the review of Emelko et al.6 Oocyst-sized fluorescent polystyrene microspheres, which have much better defined properties than living oocysts, are sometimes used as surrogates for oocysts in filtration studies.6−11 Fluorescent polystyrene microspheres are nongenotoxic12 and have often © 2012 American Chemical Society

been used as safe surrogates for microbial particles in groundwater field experiments.13,14 However, microspheres are of limited use because their surface properties (e.g., surface charge) are very different from those of the oocysts, resulting in different attachment and filtration characteristics.9,10,14 Given the variability of the surface properties of oocysts (depending on their age, isolates, strains, etc.) and differeing geologic media from site to site, microspheres can potentially over- or underpredict attenuation and transport of C. parvum oocysts.15 At the same solution chemistry, carboxylated microspheres are more negatively charged than oocysts.8 In near-neutral pH conditions, most clay and quartz minerals carry a net negative charge, but many metal-oxides, calcite, hydroxyapatite, and activated carbon carry a net positive charge.16 Carboxylated polystyrene microspheres were found to significantly deviate from oocysts removal, for example being underpredicted in silica beads10 but overpredicted in intact limestone cores.14 The differential attenuation between microspheres and oocysts is expected to be more significant under field scale. Although field data are unavailable for comparing microspheres and oocysts, we can relate this to field data that show the differential attenuation Received: Revised: Accepted: Published: 11779

June 26, 2012 September 11, 2012 September 14, 2012 September 14, 2012 dx.doi.org/10.1021/es302555n | Environ. Sci. Technol. 2012, 46, 11779−11787

Environmental Science & Technology

Article

receiving high doses of the vitamin.23 Biotin is a highly polar molecule, relatively stable, and water-soluble, with a reported IP value of 3.5.24 Two different biotin products were purchased: amine-free biotin from Sigma-Aldrich (Auckland, New Zealand) and amine-containing biotin (NH2-biotin) from Thermo Fisher Scientific Inc. (Rockford, IL). According to the suppliers, the biotin of Sigma-Aldrich had a molecular weight of 244.31 g/mol and solubility of 22 mg/100 mL, while the NH2-biotin of Thermo Fisher Scientific Inc. had a molecular weight of 374.5 g/mol and a maximum solubility of 25 mg/mL. The lyophilized powder of biotins was stored at 4 °C. Alpha 1-acid glycoprotein was purchased from Sigma-Aldrich (Auckland, New Zealand). According to the product information sheet, it has a molecular weight of 33 000−40 800 g/mol and IP = 2.7 and is highly soluble in water (solubility 1000 mg/100 mL). The α1-acid glycoprotein contains approximately 11−12% sialic acid, 13−17% neutral hexoses, 12−15% hexosamine, and 0.7−1.5% fucose (approximately 45% total carbohydrate content). Supplier’s specifications stated that no component of this product, present at levels ≥0.1%, is identified as a probable, possible, or confirmed human carcinogen. No data are available for its ecotoxicity. The lyophilized powder of glycoprotein was stored at 4 °C. 2.2. Conjugation of Biomolecules with Microspheres. Glycoprotein-coated microspheres were not commerically available for any particle sizes. Although there were some commerical biotin-coated microspheres for biochemistry applications, they were not avilable in the size range of the oocysts (4−6 μm). Thus, biotin- and glycoprotein-coated microspheres were produced in this study using the 4.87-μm microspheres. A water-soluble carbodiimide cross-linker, EDC (1-ethyl-3-(3-dimethylaminopropyl) carbodiimide hydrochloride), was used for coupling the selected biomolecules to the microspheres. EDC (C8H17N3−HCL, molecular weight 191.7 g/mol) is a versatile coupling agent to cross-link proteins and nucleic acids or to bind molecules to surfaces in aqueous or organic media. Most uses of EDC are for covalent coupling as EDC activates carboxyl groups for spontaneous reaction with primary amines. Thus the use of EDC would yield a covalent bonding of NH2-biotin and glycoprotein onto the microspheres. However the use of EDC for amine-free biotin would result in adsorption rather than covalent bonding. The coupling procedures of Bangs Laboratories, Inc.25 were followed to conjugate the selected biomolecules onto microsphere surfaces. A detailed description of the procedures is given elsewhere.25 Washes and separations were performed using centrifugation. During the coupling, physical methods of rolling, rotation, vortexing, and sonication were used to keep the modified microspheres in a monodispersed state. The modified beads were stored in phosphate buffer saline (pH 7.4) with bovine serum albumin blocker (0.05%). The solutions were then stored at 4 °C until use. 2.3. Determination of Zeta Potentials and Charge Stability. The zeta potentials of the C. parvum oocysts (within two weeks of receiving the stock), biomolecules, and modified and unmodified microspheres were determined using laser Doppler microelectrophoresis (Malvern Instruments Zetasizer Nano ZS). Prior to the zeta potential measurement, each sample was sonicated in a cold water bath for 3 min. The zeta potentials (ζ) were measured in a background electrolyte of 1 mM NaCl at various pH values (adjusted with 10 mM NaOH

between bacteria and bacterium-sized microspheres. By analyzing previous field data, Harvey et al.13 showed that in comparison with bacteria, the fractional losses of microspheres were 1.5−2.4 log units greater in the Cape Cod sand aquifers and 0.5−1.4 log units greater in limestone and fractured granite aquifers, but 0.5−1.4 log units lower in a granite aquifer for microbacterium strain. Surface charge plays a critical role in particle attachment onto solid surfaces17,18 and filtration.19 Thus for microspheres to be a better oocyst surrogate and to give more accurate predictions, their surface charges need to be modified to approximate that of the oocysts. Pang et al.20 proposed mimicking the surface charges of a microorganism using microspheres coupled with a biomolecule that had surface charge characteristics similar to the microorganism. They demonstrated that when the 20-nm carboxylated polystyrene nanospheres were coupled with bovine milke αs-casein, which had an isoelectric point (IP) similar to that of MS2, the modified nanospheres displayed a surface charge similar to that of MS2. However, they did not validate the resemblance of the modified nanospheres to MS2 in their attenuation and transport behaviors because of a difficulty in detecting the nanospheres. To the best of our knowledge, no study has been reported in mimicking pathogen attenuation and transport in porous media using biomoleculemodified microspheres. In this study, the principle of Pang et al.20 was applied to develop surrogates for C. parvum by mimicking its size, surface charge, shape, and density, and if possible, macromolecular structure. Column filtration experiments with sands of two different grain sizes were carried out to examine whether the surrogates would better represent concentration reduction of the oocysts during filtration process than traditional unmodified microspheres.

2. MATERIALS AND METHODS 2.1. Cryptosporidium parvum, Microspheres, and Biomolecules. C. parvum oocysts are spherical or oval, 3.9− 5.9 μm in diameter,21 and have an IP ≈ 2.5.22 A suspension of live unpreserved C. parvum was purchased from Waterborne, Inc. (New Orleans, LA). No inactivation was carried out on the oocysts prior to the experiments, and the oocysts viability was not assessed. The oocysts were stored in phosphate buffered saline with antibiotics and 0.01% Tween 20 in the dark at 4 °C. The suspension was used in the experiments within 5 months of receiving the stock. To mimic the size, shape, and density of the oocysts, oocystsized carboxylated polystyrene microspheres were purchased from Polysciences, Inc. (Warrington, USA). The microspheres had a mean diameter of 4.87 μm (standard deviation SD = 0.12) as provided by the manufacturer and were bright blue (emission 407 nm, excitation 360 nm) with a concentration of 4.99 × 108 particles/mL. The manufacturer’s specifications stated that the buoyant density of polystyrene microspheres was 1.05 g/cm3, which equals the geometric mean of the oocyst density.21 The microsphere suspension was stored in the dark at 4 °C. To mimic the surface charge of the oocysts, biotin and glycoprotein, with IP values similar to that of the C. parvum oocysts, were selected. Biotin (C10H16N2O3S) is B-complex vitamin (vitamin B7) found in a wide range of foods (e.g., yeast, liver, kidney, egg yolk, soybeans, nuts, and cereals). Animal studies have indicated few, if any, effects due to toxic doses of biotin, and there are no reported cases of adverse effects from 11780

dx.doi.org/10.1021/es302555n | Environ. Sci. Technol. 2012, 46, 11779−11787


Article

Table 1. Parameter Values of Sand Columns Used in the Filtration Study column information

CXTFIT-optimized from Br data

IDa

runb

porosity

Uc (cm/min)

Dc (cm2/min)

r2d

D50 = 0.78 mm sand uncoated microspheres

A

noninactivated C. parvum

B

glycoprotein-coated microspheres

C

NH2-biotin-coated microspheres

D

biotin-coated microspheres

E

1 2 1 2 1 2 1 2 1 2

0.41 0.41 0.41 0.41 0.42 0.42 0.49 0.49 0.41 0.41

0.07 0.06 0.06 0.06 0.05 0.06 0.06 0.06 0.07 0.07

0.02 0.02 0.04 0.04 0.01 0.01 0.01 0.01 0.02 0.02

1.00 1.00 1.00 0.99 0.98 0.96 0.96 0.96 0.96 0.98

1 1 1 2 1 2 1

0.38 0.38 0.38 0.38 0.42 0.42 0.38

0.07 0.07 0.07 0.06 0.05 0.06 0.07

0.06 0.03 0.03 0.06 0.02 0.02 0.06

0.99 0.99 1.00 0.98 1.00 1.00 0.99

experiment

D50 = 1.37 mm sand uncoated microspheres noninactivated C. parvum

F G H

glycoprotein-coated microspheres

I

biotin-coated microspheres

J

particle data measured

particle data calculated

C0e (particles/mL)

log10 (max C/C0)

RBf (%)

αg

5.00 5.00 3.90 6.06 5.70 6.67 5.73 5.73 5.00 5.00

× × × × × × × × × ×

106i 106i 104h 104h 106i 106i 106i 106i 106i 106i

1.6 1.6 2.7 3.0 2.6 3.4 2.4 2.6 2.5 2.6

9.1 6.0 1.6 0.6 3.7 0.7 5.1 1.6 1.0 0.9

0.03 0.03 0.05 0.06 0.03 0.05 0.04 0.05 0.06 0.06

2.80 5.80 1.00 5.00 6.77 6.77 2.80

× × × × × × ×

105h 104h 103h 104h 106i 106i 105h

1.5 1.5 2.3 2.3 2.0 2.0 2.1

4.4 5.5 1.6 0.8 5.5 3.1 1.0

0.06 0.06 0.09 0.08 0.05 0.06 0.09

ID: Column identification number. bRun 1: Column packed with clean sand, Run 2: Column flushed after Run 1. cDarcy velocity (U = pore velocity × porosity) and dispersion coefficient (D) were estimated from CXTFIT modeling of Br breakthrough data. dr2: Goodness of fit from CXTFIT modeling. eC0: Input concentration. fRB: Normalized mass recovery. gα: Collision efficiency. hCounted by epifluorescence microscope. i Counted by spectrofluorometer. a

elsewhere.27 Prior to the measurements, microspheres were sonicated in a sonicating water bath for 5 min and the solution was diluted in 20 mM NaCl at pH = 7. 2.5. Filtration Column Experiments. Glass columns (22 cm long and 5 cm in diameter) were packed under saturated condition with coarse sand (d50 = 0.78 mm, d60/d10 = 2.36, where subscript is the percentile in a cumulative frequency curve of particle size) and very coarse sand (d50 = 1.37 mm, d60/d10 = 1.94), respectively. These materials were natural sand sourced from alluvial deposits in the Canterbury Plains (New Zealand). Prior to packing into the columns, the sand materials were washed using tap water followed by rinsing in 1 mM NaCl solution and oven-dried. Different types of particles (oocysts and unmodified and modified microspheres) were examined independently in columns repacked with clean sand. For each type of particles, there were two mostly sequential runs in the same column, except for the uncoated and biotin-coated microspheres in 1.37-mm sand. After the first run, the column was flushed with tracer-free 1 mM NaCl solution (pH = 7) until particle concentrations were reduced to the background levels. There were five freshly packed columns for the 0.78-mm sand and five freshly packed columns for the 1.37-mm sand, producing a total of 17 experiments (Table 1). In each experiment a pulse of tracer solution (0.11−0.15 pore volume) containing one type of test particles and bromide (Br 0.15−0.35 mM) was injected. Br was used to indicate flow. Input particle concentrations for most experiments were 106 microspheres/mL and 104 oocysts/mL (Table 1). The difference in input concentrations between microspheres and oocysts was because of the different detection limits between spectrofluorometry and microscopy (section 2.6). Exceptions were that in the four earlier experiments with 1.37-mm sand, lower input concentrations (104−105 microspheres/mL, 103 oocysts/mL, Table 1) were used. This is because in these

or HCl). The measurements were carried out for 3−5 replicates of each pH. There was concern that the charge modification using NH2free biotin by adsorption may not be permanent because the NH2-free biotin was not covalently coupled to the microspheres. To ascertain this, the zeta potentials of biotin-coated microspheres were re-examined 21 months after the couplings at pH 6.0, 6.5, 7.0, and 7.5 in 1 mM NaCl. A short-term monitoring of zeta potentials was carried out for glycoproteincoated microspheres over 19 days. Long-term monitoring of charge stability was not carried out for NH2-biotin- and glycoprotein-coated microspheres because the stocks ran out after the column experiments. To examine the ζ of sand media used in the column study, sand samples were crushed to colloid-sized powder so that their suspensions could be measured by the Zetasizer, adapting the method of Sharma et al.26 A large Rocklabs (TEMA) swing mill with a tungsten carbide mill head was used with 1 min crushing for each sample. The mill head was washed thoroughly between each sample. Approximately 10 mg of the resulting powder was then dispersed in 15 mL of 1 mM NaCl. The suspensions were sonicated for 10 min and their ζ measured at pH = 7.0 in triplicates, giving ζ = −29.6(±0.74) mV for 0.78-mm sand and ζ = −23.1(±0.10) mV for 1.37-mm sand. 2.4. Particle Characterization. The modification of microsphere surfaces may result in a change in particle size and a loss in particle concentration during washing procedure. To ascertain this, a newly developed qNano Analyzer (V1.0 model, IZON Science, Christchurch, New Zealand) was used for particle characterization for the microspheres and oocysts. The qNano Analyzer characterizes particle sizes, size distribution (ranging 40 nm−10 μm), particle concentration, and relative surface charge distributions. qNano’s working principle and detailed experimental setup have been reported 11781



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where C/C0 is the relative concentration (dimensionless), t is the time since injection (min), and t0 and tf are the times of the beginning and end of the breakthrough (min). Using the normalized mass recovery has advantages over using the actual mass recovery when BTCs produced from different experiments are compared. When Br reaches 100% mass recovey, the particle RB value equals the actual particle mass recovery. However, when Br mass recovery is less than 100% or BTCs are incomplete, it would be inaccurate to directly compare actual particle mass recoveries from different experiments but using normalized mass recovery allows a direct comparison. The collision efficiency, α (the ratio of the rate of collisions resulting in attachment to the total rate of collisions between particles and collector grains), was calculated using the following formula:29

earlier experiments microsphere samples were also enumerated using the epifluorescence microscope (section 2.6). Prior to each experiment, the stock was sonicated in a sonicating water bath for 5 min to detach any temporarily bound particle−particle aggregates. Flow was introduced using a peristaltic MilliGAT Pump (Global FIA, Inc.) at the bottom of the column to ensure that trapped air was removed and the wall effect was minimized. A constant flow rate (at an equivalent Darcy velocity of 0.8−1.0 m/day) was applied. During each experiment, flow rate was regularly measured. To maintain the same water chemistry, the same background electrolyte of 1 mM NaCl at pH ≈ 7 was used in all the experiments. The condition of ionic strength 1 mM and pH ≈ 7 was similar to that of the tap water (sourced from groundwater) in Christchurch city (New Zealand). All experiments were carried out at room temperature (∼18 °C). Samples of the injection solution and column outflow were taken. The Br samples were analyzed using a bromide ion selective electrode. Microspheres and oocysts were prepared and assayed using the method described below. 2.6. Enumeration of Particles. C. parvum oocysts were enumerated using U.S. EPA Method 1623.28 The oocyst samples were filtered through black nucleopore filters (0.22 μm pore size, Whatman). The filter was stained with fluorescein isothiocyanate (FITC) stain (Crypt-a-Glo kit, Waterborne Inc., USA) and incubated in a humid chamber at 37 °C for 30 min. Microscopic examinations were performed using an epifluorescence microscope (Leica instruments) with excitation/ bandpass filter for FITC (450−490 bp). Each filter was scanned at a magnification of ×400 and the number of oocysts present was counted. The microscope had a detection limit of 1 particle/mL. Microsphere samples from the first three experiments (unmodified and biotin-coated microspheres with 1.37-mm sand) were enumerated using the same epifluorescence microscope with excitation/Bandpass filter for UV (360/40 bp). Each filter was scanned at a magnification of ×100 and the number of microspheres present was counted. A comparison of microsphere concentration between microscopy and spectrofluorometry gave similar results. Thus, for the remaining experiments, all the microsphere samples were analyzed by spectrofluorometry. The spectrofluorometer used was a FP6300 model (JASCO International Co., Ltd.) with emission and excitation wavelengths of 407 and 360 nm, respectively. The spectrofluorometer had a detection limit of 103 particles/mL for the bright blue microspheres used. There was no interference from background fluorescence on the detection. 2.7. Evaluation of Filtration Removal. Filtration removal of particles was measured using peak breaththrough attenuation, normalized mass recovery, and collision coefficient. The peak breakthrough attenuation is the log-reduction of the peak effluent concentration (Cmax) relative to the input concentration (C0), log(Cmax/C0). The normalized mass recovery (RB)29 was calculated from the zero moments of the particles and Br by inegrating their breathroough curves (BTC) ising the trapezoidal rule

α=

t0 tf 0

2

)

6(1 − θ )ηλ

⎤ − 1⎥ ⎦ (2)

3. RESULTS AND DISCUSSION 3.1. Surface Charge Modifications and Charge Stability. Figure 1a shows that the zeta potentials of biotin, glycoprotein, and noninactivated C. parvum oocysts were very similar with IP ≈ 2. Biotin and glycoprotein were slightly more negatively charged than oocysts, especially at pH 5−9. The value of IP ≈ 2 for C. parvum was very similar to IP = 2.5 reported by Butkus et al.22 In contrast, the unmodified microspheres exhibited significantly greater negative charges than the oocysts. They remained negatively charged throughout the entire pH range studied (pH = 2−10). A much greater negative charge on carboxylated microspheres than on oocysts was also reported by others.8,9,14,34 The zeta potentials of the microspheres changed significantly after being coated with biotin, NH2-biotin, and glycoprotein (Figure 1b), becoming less charged than the unmodified microspheres and more similar to those of the oocysts. The glycoprotein-coated microspheres best matched the zeta potentials of C. parvum, while NH2-free biotin provided the least match. A better ζ result from NH2-biotin than from biotin was expected as covalent bonding is much stronger than adsorption. Despite the fact that the amine-free biotin was not covalently coupled to the microspheres, the modified surface charge was very stable over 22 months. This was demonstrated in the similar ζ results measured at month 0 and month 22 (Figure 2a). A stable charge was also observed for a short-term (19

tf

∫ [(C(t )/C0)dt ]Br

(

where d is the mean grain size of the porous media (mm) using d50, λ is the dispersivity (mm), x is the column length (mm), θ is the effective porosity, and η is the single-collector efficiency. The η value was calculated based on the modified equations published by Tufenkji and Elimelech,30 using an Excel spreadsheet provided from the authors’ webpage (http:// www.yale.edu/env/elimelech/publication-pdf/ TECorrelationEqn.xls).31 A Hamaker constant of 1.00 × 10−20 J for glass−water−polystyrene32 and 6.50 × 10−21 J for oocysts33 were used. As Darcy velocity (U, in mm/min and dispersivity (λ in mm) are required in the calculations of η and α, Br data were simulated using the CXTFIT transport model to derive pore-velocity (V = U/θ, in mm/min) and dispersion coefficient (D = λV, in mm2/min) by fitting the advection− dispersion equation.

∫ [(C(t )/C0)dt ]particles RB =

⎡ λ d⎢ 1 − 2 x ln(RB) ⎣

(1) 11782



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Figure 1. Zeta potentials of particles and biomolecules measured in 1 mM NaCl electrolyte: (a) noninactivated C. parvum (red ●), biotin (green ▲), glycoprotein (brown ▼), and unmodified microspheres (blue ■) and (b) microspheres coated with biotin (green ▲), NH2biotin (dark green ⧫), and glycoprotein (brown ▼), noninactivated C. parvum (red ●), and unmodified microspheres (blue ■). Average and one standard deviation of 3−5 replicate measurements are shown.

Figure 2. Stability of zeta potentials for (a) biotin-coated microspheres at 0 (○) and 22 (●) months after coating at a range of pH and (b) glycoprotein-coated microspheres over 19 days at pH 7. Average and one standard deviation of the 3−5 replicate measurements are shown.

Table 2. Geometric Mean of Particle Size and Uniformity Measured from qNano

days) monitoring of glycoprotein-coated microspheres (Figure 2b). Although long-term monitoring of charge stability was not carried out for microspheres coated with glycoprotein and NH2-biotin, their surface charges were expected to be stable because covalent coupling produces permanent bonding. For example, Pang et al.20 observed essentially unchanged IP values and the pH−ζ patterns (pH = 3−9) for nanospheres covalently bound to αs-casein over a period of four months. 3.2. Particle Characterization. The results of qNano Analyzer measurements (Table 2) suggest that the mean sizes of unmodified and modified microspheres were very similar to that of the oocysts ∼4.9 μm. This indicates that surface coating had not significantly changed the particle size of the microspheres. The mean size of 4.86 (±0.24) μm for C. parvum oocysts determined in this study was very similar to 4.9 (±0.3) μm value previously reported.21 Likewise, the mean size of 4.92 (±0.03) μm (summarized from 3 samples) for unmodified microspheres was very similar to 4.87 (±0.12) μm stated by the supplier. Table 2 also shows that the uniformity of particle size distribution for unmodified and modified microspheres, as indicated from ϕ90/ϕ10 (subscript is the percentile in a cumulative frequency curve of particle size), was also very similar to that of the oocysts. A ϕ90/ϕ10 value of 1.2 suggests

sample noninactivated C. parvum unmodified microspheres glycoprotein-coated microspheres biotin-coated microspheres NH2-biotin-coated microspheres

ϕa mean (μm)

SDb (μm)

ϕ90/ ϕ10c

SD

no. of samples

4.86

0.24

1.20

0.06

1

4.92

0.03

1.23

0.06

3

4.95

0.18

1.20

0.00

3

4.93

0.17

1.15

0.07

2

4.88

0.09

1.15

0.07

2

ϕ: Diameter of particles. bSD: Standard deviation. cϕ90 and ϕ10 are the 90 and 10 percentile in a cumulative frequency curve of particle size.

a

that particle size distribution was relatively uniform and these particles were monodispersed in solutions. This indicates that sonication could effectively detach temporarily bound particle− particle aggregates. The qNano-measured oocyst concentration (2 × 10 6 particles/mL) was in the same order of magnitude as given by the supplier (6 × 106 particles/mL). qNano results also showed that concentrations of modified microspheres were slightly lower than the unmodified microspheres but the 11783



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difference was still within 1 order of magnitude. A loss of some microspheres during washing procedures of coupling was expected. 3.3. Filtration Removal. Figures 3 and 4 clearly demonstrate that, in comparison with the unmodified micro-

Figure 4. BTCs of noninactivated C. parvum (red ●), unmodified (blue ■), and microspheres coated with glycoprotein (brown ▼) and biotin (green ▲) from 1.37-mm sand columns (ID F, G, H, I, J) in 1 mM NaCl electrolyte (a) columns packed with clean sand and (b) flushed columns. For C. parvum and glycoprotein-coated microspheres, two experiments were carried out sequentially in the same column. Unmodified microspheres in (b) were from a column packed with clean sand. BTCs of 7 experiments are shown and each BTC was produced from an independent experiment (refer to Table 1).

effect of input concentration on particle attenuation seemed not very sensitive for the concentration range investigated. Like the oocysts, BTCs of the modified microspheres showed marked variability, while those of the unmodified microspheres were much smoother and contained significantly less variability (Figure 3a). This was possibly because of the extra surface structure introduced from the coating. Of the three types of modified microspheres, glycoproteincoated microspheres showed the best match to oocyst peak concentrations and tailings. Microspheres coated with biotin and NH2-biotin slightly overpredicted oocysts peak concentrations, and the biotin-coated microspheres underpredicted the tailings. Although the zeta potentials of glycoprotein-coated and NH2-biotin-coated microspheres were very similar (Figure 1b), glycoprotein-coated microspheres better mimicked the macromolecule structure of the oocysts. This is because C. parvum possesses numerous glycoproteins on its surfaces35,36 and has an acidic glycoprotein outer layer on oocyst walls.37 Our findings suggest that surface protein can play an important role in oocyst attachment to solid surfaces. It has been found that macromolecules play a significant role in the attachment of C. parvum oocysts to solid surfaces.38 Kuznar and Elimelech33 observed that despite having near identical zeta potentials, inactivated (with heat or formalin) oocysts displayed significantly greater attachment efficiency than viable oocysts

Figure 3. BTCs of noninactivated C. parvum (red ●), unmodified (blue ■), and microspheres coated with biotin (green ▲), NH2-biotin (dark green ⧫), glycoprotein (brown ▼) from 0.78-mm sand columns (ID A, B, C, D, E) in 1 mM NaCl electrolyte (a) columns packed with clean sand and (b) flushed columns. Inset graphs with the expanded scale show all data except for unmodified microspheres. For each type of particle, two experiments were carried out sequentially in the same column. BTCs of 10 experiments are shown and each BTC was produced from an independent experiment (refer to Table 1).

spheres, the modified microspheres mimicked more closely the concentrations of oocysts for both sand media either with or without repacking the columns with clean sand. Typically the modified microspheres showed the same magnitude of concentration recovery as that of oocysts. In contrast, those of the unmodified microspheres deviated from the oocysts by 1 order of magnitude (Table 1). For 0.78-mm sand, reduction in peak concentration was ∼3-log for the oocysts and modified microspheres but ∼2-log for the unmodified microspheres. For 1.37-mm sand, the concentration reduction was 2.0−2.3-log for the oocysts and glycoprotein-coated microspheres but 1.5-log for the unmodified microspheres. This pattern was consistent despite the variations in input concentrations (Table 1). The 11784



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concentrations could save unnecessary treatment costs than if predicted using the overly conservative unmodified microspheres. Although their concentration difference was about 1 order of magnitude for the sand media investigated here, this difference may vary in other porous media. For field scale operations treatment costs may increase significantly if treatment level required is 1-log higher. Therefore the use of a surrogate such as glycoprotein-coated microspheres, which could more accurately predict oocyst removal than unmodified microspheres, is more appropriate. The work presented here was from a laboratory study carried out in simple experimental conditions and a background solution of low ionic strength at near neutral pH. Future work is needed to evaluate the usefulness of the surrogates developed here in environmental conditions, for example with high organic matter content, a range of pH conditions, high ionic strength, oocysts of different surface properties (e.g., age, isolates), and in the presence of divalent ions (e.g., Ca2+ and Mg2+) and naturally occurring bacteria. Although polystyren microspheres may hold promise as a surrogate for Cryptosporidium removal by filtration,6 little has been reported on the impact of accumulated polystyrene microspheres under environmental conditions. This is another area of future study. Significant costs can be save by using easily detectable surrogates. Fluorescently tagged microspheres can be rapidly enumerated using user-friendly automatic countring techniques (e.g., a spectrofluorimeter, flow cytomerter, or particle counter. The result can be delivered wthin a few minutes per sample in a one-step process in a non-microbiology laboratory. In contrast, enumeration of the oocysts could take a few hours for processing (involving multiple steps) and analyzing one environmental sample. Dealing with pathogenic samples requires a PC2-compliant microbiology laboratory (with epiflourescence microscope) and specially trained technical staff. The costs of C. parvum culture and consumables for processing the samples are also expensive compared to the use of microspheres. C. parvum cannot be grown because they must pass through animals. In contrast, synthesis cost of the miscrosphere surrogates can be significantly reduced as volume increases. Although further validation is needed, our study has nevertheless made good progress in the development of better surrogates for quantifying oocyst removal by filtration. With further evaluation and pilot studies, the surrogates developed in this study could be a cost-effective new tool for predicting filtration removal of C. parvum oocysts in porous media (especially those containing macropores), for example evaluation of filter performance in water and wastewater treatment, river bank filtration, and aquifer passage and recharge.

to quartz surfaces. They attributed this to a disruption of surface proteins during the inactivation. We expect that this will also apply for the UV inactivated oocysts because of protein denaturation under the UV light.39 For 0.78-mm sand columns, when the same column was used for two sequential experimental runs, peak concentration of the oocysts or modified microspheres was notably reduced in the second run, resulting in an increased log-reduction (Table 1, Figure 3b). However there was no significant difference for unmodified microspheres. For 1.37-mm sand columns, there was no significant change in peak concentrations of particles between the two sequential runs. The size ratio of oocysts and microspheres to sand grains (ϕ/d50) was 0.63% for 0.78-mm sand and 0.36% for 1.37-mm sand. Straining occurs when the ratio of the colloid to medium grain diameter is >0.5%.9 Thus the above observations may be explained by the presence of straining in 0.78-mm sand and its absence in 1.37-mm sand. Compared to unmodified microspheres, the modified microspheres also showed a closer mimic to mass recovery of the oocysts in 0.78-mm sand, as suggested in the RB values. In contrast, the unmodified microspheres overpredicted mass recoveries of the oocysts because their breakthroughs were significantly greater than those of the oocysts. For all the tested particles, mass recoveries were lower in the flushed columns than in the freshly packed columns. This was because particles retained in the columns from the first experimental runs would have acted as additional attachment sites resulting in a reduced filtration removal. Mass recoveries of the oocysts were very similar in both 0.78-mm and 1.37-mm sand, being ∼2% in the freshly packed columns and ∼1% in the the flushed columns. Likewise, mass recoveries of biotin-coated microspheres were very similar in two sand media (∼1%). However, glycoproteincoated microspheres recovered 2% more in the 1.37-mm sand than in the 0.78-mm sand. The degree of oocyst removal (∼2 logs removal in mass and 2−3 logs in peak breakthrough attenuation)in sand media observed in this study was within the range of oocyst removal in sand media (2−5 logs) as summarized in a review.6 Compared to the measured concentrations and RB values, the calculated α values contained a greater level of uncertainty. Many parameters were involved in the calculations, and each parameter value had its own uncertainty and variation. Thus we did not carry out further modeling using a kinetic transport model to determine attachment and detachment rates of the particles. Nevertheless, the modified microspheres gave an overall better prediction of α values of the oocysts than did the unmodified microspheres (Table 1). The unmodified microspheres mostly underpredicted the α values of the oocysts. 3.4. Implications and Future Research. Although the unmodified microspheres had less retention than the oocysts in the sand media investigated in this study because they were more negatively charged, unmodified microspheres will not necessarily be conservative in all types of porous media. The possibility of greater removal exists for unmodified microspheres than for the oocysts in the presence of positively charged media. For example, metal (hydr)oxide-coated sand filters have been developed for effective removal of microorganisms and other contaminants in water and wastewater treatment.40 Therefore using unmodified microspheres in such a treatment plant may overpredict the ability of the system to remove oocysts. If the filtration media used is net-negatively charged, the use of a surrogate that will more accurately predict oocyst

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AUTHOR INFORMATION

Corresponding Author

*E-mail: [email protected]. Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS We thank Julianna Isgar (Bangs Laboratories, Inc.) for her valuable comments on surface coupling, and Murray Close, Jacqui Horswell, Chris Nokes, and Wendy Williamson (Institute of Environmental Science and Research Ltd.) for their helpful comments on the manuscript. This study was 11785



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(19) Emelko, M. B. Removal of viable and inactivated Cryptosporidium by dual- and tri- media filtration. Water Res. 2003, 37 (12), 2998−3008. (20) Pang, L.; Nowostawska, U.; Ryan, J. N.; Williamson, W. M.; Walshe, G.; Hunter, K. A. Modifying the surface charge of pathogensized microspheres for studying pathogen transport in groundwater. J. Environ. Qual. 2009, 38 (6), 2210−2217. (21) Medema, G. J.; Schets, F. M.; Teunis, P. F. M.; Havelaar, A. H. Sedimentation of free and attached Cryptosporidium oocysts and Giardia cysts in water. Appl. Environ. Microbiol. 1998, 64 (11), 4460− 4466. (22) Butkus, M. A.; Bays, J. T.; Labare, M. P. Influence of surface characteristics on the stability of Cryptosporidium parvum oocysts. Appl. Environ. Microbiol. 2003, 69 (7), 3819−3825. (23) Combs, G. F. J. The Vitamins: Fundamental Aspects in Nutrition and Health; Academic Press: San Diego, CA, 1998. (24) Khamaisi, B.; Vaknin, O.; Shaya, O.; Ashkenasy, N. Electrical performance of silicon-on-insulator field-effect transistors with multiple top-gate organic layers in electrolyte solution. ACS Nano 2010, 4 (8), 4601−4608. (25) TechNote 205Covalent Coupling; Bangs Laboratories Inc.: Fishers, IN, 2008; Vol. KT/MM-03/2002, p 11; http://www. bangslabs.com/sites/default/files/bangs/docs/pdf/205.pdf. (26) Sharma, P.; Flury, M.; Zhou, J. Detachment of colloids from a solid surface by a moving air−water interface. J. Colloid Interface Sci. 2008, 326 (1), 143−150. (27) Vogel, R.; Willmott, G.; Kozak, D.; Roberts, G.; Anderson, W.; Groenewegen, L.; Glossop, B.; Barnett, A.; Turner, A.; Trau, M. Quantitative sizing of nano/microparticles with a tunable elastomeric pore sensor. Anal. Chem. 2011, 83 (9), 3499−3506. (28) U.S. Environmental Protection Agency. Method 1623: Cryptosporidium and Giardia in Water by Filtration/IMS/FA; U.S. EPA: Washington DC, 2005; EPS 815-R-05002, p 76. (29) Pieper, A. P.; Ryan, J. N.; Harvey, R. W.; Amy, G. L.; Illangasekare, T. H.; Metge, D. W. Transport and recovery of bacteriophage PRD1 in a sand and gravel aquifer: Effect of sewagederived organic matter. Environ. Sci. Technol. 1997, 31 (4), 1163− 1170. (30) Tufenkji, N.; Elimelech, M. Correlation equation for predicting single-collector efficiency in physicochemical filtration in saturated porous media. Environ. Sci. Technol. 2003, 38 (2), 529−536. (31) Tufenkji, N.; Miller, G. F.; Ryan, J. N.; Harvey, R. W.; Elimelech, M. Transport of Cryptosporidium oocysts in porous media: Role of straining and physicochemical filtration. Environ. Sci. Technol. 2004, 38 (22), 5932−5938. (32) Elimelech, M.; O’Melia, C. R. Kinetics of deposition of colloidal particles in porous media. Environ. Sci. Technol. 1990, 24 (10), 1528− 1536. (33) Kuznar, Z. A.; Elimelech, M. Role of surface proteins in the deposition kinetics of Cryptosporidium parvum oocysts. Langmuir 2005, 21 (2), 710−716. (34) Tufenkji, N.; Elimelech, M. Spatial distributions of Cryptosporidium oocysts in porous media: Evidence for dual mode deposition. Environ. Sci. Technol. 2005, 39 (10), 3620−3629. (35) Cevallos, A. M.; Zhang, X.; Waldor, M. K.; Jaison, S.; Zhou, X.; Tzipori, S.; Neutra, M. R.; Ward, H. D. Molecular cloning and expression of a gene encoding Cryptosporidium parvum glycoproteins gp40 and gp15. Infect. Immun. 2000, 68 (7), 4108−4116. (36) Waldron, L. S.; Ferrari, B. C.; Power, M. L. Glycoprotein 60 diversity in C. hominis and C. parvum causing human cryptosporidiosis in NSW, Australia. Exp. Parasitol. 2009, 122 (2), 124−127. (37) Harris, J. R.; Petry, F. Cryptosporidium parvum: Structural components of the oocyst wall. J. Parasitol 1999, 85 (5), 839−849. (38) Kuznar, Z. A.; Elimelech, M. Cryptosporidium oocyst surface macromolecules significantly hinder oocyst attachment. Environ. Sci. Technol. 2006, 40 (6), 1837−1842. (39) Vladimirov, Yu. A; Roshchupkin, D. I.; Fesenko, E. E. Photochemical reactions in amino acid residues and inactivation of

funded by the Foundation for Research, Science and Technology (New Zealand).

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Biotin- and Glycoprotein-Coated Microspheres: Potential Surrogates

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